The effects of earthworms Eisenia spp. on microbial community are habitat dependent

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Original article
The effects of earthworms Eisenia spp. on microbial community are
habitat dependent
Anna Koubov
a
a
,
b
,
*
, Alica Chro
n
akov
a
a
,V

aclav Pi
zl
a
, Miguel Angel S
anchez-Monedero
c
,
Dana Elhottov
a
a
a
Institute of Soil Biology, Biology Centre CAS, Na S
adk
ach 7, CZ-37005 
Cesk
e Bud
ejovice, Czech Republic
b
Faculty of Science, University of South Bohemia, Brani
sovsk
a 31, CZ-37005 
Cesk
e Bud
ejovice, Czech Republic
c
Department of Soil and Water Conservation and Organic Waste Management, Centro de Edafología y Biología Aplicada del Segura, CSIC, P.O. Box 4195, E-
30080 Murcia, Spain
article info
Article history:
Received 10 November 2014
Received in revised form
8 March 2015
Accepted 11 March 2015
Available online 13 March 2015
Keywords:
Earthworms
Soil
Compost
Vermiculture
Archaea
Bacteria
abstract
The effects of earthworms Eisenia spp. on microorganisms of three different habitat soil, compost, and
vermiculture were studied. Microbial communities of gut and fresh faeces of earthworms and substrates,
the worms were collected from, were analysed. Microbial biomass and composition of the total microbial
community were examined using phospholipid fatty acid (PLFA) biomarkers. Archaeal and bacterial
communities were studied by polymerase chain reaction-denaturing gradient gel electrophoresis
(DGGE). The culturing methods were used for assessment of counts, species richness and growth strategy
of bacteria.
In comparison with the substrates, the viable microbial biomass and the group of non-ester-linked
PLFAs indicative of anaerobes were higher in both the gut and faeces of all earthworm populations.
The prokaryotic community evaluated using DGGE revealed that archaeal community structure in the gut
and faeces of earthworms from populations differed from that in substrates, whereas the passage
through the gut had less influence on the bacterial community structure, particularly in compost and
vermiculture.
The counts of culturable bacteria increased due to gut passage only in forest and vermiculture pop-
ulations. The fast-growing bacteria increased due to gut passage only in forest soil population. Actino-
bacteria (Arthrobacter,Microbacterium,Lechevalieria and Nesterenkonia) and Firmicutes (Bacillus and
Paenibacillus) were generally favoured in substrates and their species richness decreased with gut pas-
sage, whereas Gammaproteobacteria (Aeromonas,Enterobacter,Pseudomonas and Salmonella) dominated
in gut contents. The impact of earthworm activity on the microbial community was higher in nutrient-
poor forest soil than in nutrient-rich compost and vermiculture substrates.
©2015 Published by Elsevier Masson SAS.
1. Introduction
Earthworms are remarkable drivers of decomposition of dead
organic matter and bioturbation processes in soil; they increase
mineralisation rate of organic matter due to the enhancement of
microbial activity [1] and modify the physical structure of soil.
Earthworms come into interactions with microorganisms not only
during their direct ingestion but also affect microbial life indirectly
by forming and altering their habitat [2]. Their casts enrich the
microbial nutrient status [3]. Passage of feeding material through
the earthworm gut leads to a different bacterial composition of gut
and faeces compared to that of the original soil or compost [46].
The shift in microbial biomass, structure and species diversity of
the intestinal microbial community has been previously studied in
different earthworm ecological groups (anecic, endogeic and
epigeic), however the data are fragmented. Sampedro et al. [6] and
Sampedro and Whalen [7] revealed that the total microbial biomass
(expressed as PLFA concentration) and biomarkers of aerobic bac-
teria, microeukaryotes and fungi increased in gut of anecic Lum-
bricus terrestris compared to bulk soil. In contrast, Marhan et al. [8]
did not found the differences between total amount of PLFA in soil
and gut of the same species. Counts of culturable bacteria isolated
*Corresponding author. Institute of Soil Biology, Biology Centre CAS, Na S
adk
ach
7, CZ-37005 
Cesk
e Bud
ejovice, Czech Republic.
E-mail address: koubova.anna@centrum.cz (A. Koubov
a).
Contents lists available at ScienceDirect
European Journal of Soil Biology
journal homepage: http://www.elsevier.com/locate/ejsobi
http://dx.doi.org/10.1016/j.ejsobi.2015.03.004
1164-5563/©2015 Published by Elsevier Masson SAS.
European Journal of Soil Biology 68 (2015) 42e55

from gut of epigeic Eisenia fetida increased compared to soil and
were found three fold higher than those from anecic (L.terrestris)
and endogeic earthworms (Aporrectodea caliginosa) that remained
constant [9,10]. In contrast to other earthworm species, relatively
little is known about the prokaryotic microorganisms (archaea and
bacteria) associated with the intestinal tract of epigeic E. fetida and
Eisenia andrei [1114] and only few studies include both species
[11,1517]. To date archaea have been studied as associated with
the intestine of Eudrilus eugeniae [18] and Lumbricus rubellus [19]
and culturable anaerobes have been estimated in L.rubellus and
Octolasion lacteum [9].
Thakuria et al. [20] found that earthworm ecological group is the
strongest factor for the composition of the gut-wall associated
bacteria. Little is known about the influence of earthworm habitat
on microbiota within the same ecological group on the microbial
changes in earthworm gut [8,21]. To study the effects of different
feeding habitats on microbiota associated with the intestine of
earthworms we tested three earthworm populations living in three
specific habitats: an indigenous soil population of E. fetida and a
compost and a vermiculture population of E. andrei, representing
the same ecological group.
The epigeic earthworms E. fetida (Savigny, 1826) and E. andrei
Bouch
e (1972) are closely related sibling species with similar
anatomy and morphology [22]. Both species can be properly
differentiated at the molecular level [16]. In nature E.fetida and E.
andrei lives in fresh litter layer of forest soil, in litter mounds and
synantropicaly in manure heaps, herbivore dungs and compost
pile. In Central Europe E.andrei commonly inhabits antropogenic
cultures, although E.fetida lives in unique forest populations [23].
Both species are used for the management of organic wastes as
vermicomposting [24], in ecotoxicology [2527], in physiology
[28], and as model organisms in immunology to understand the
defense mechanisms of soil invertebrates again microbial patho-
gens [2932].
The aim of this study was to determine (i) the microbial changes
occurring during the passage of the substrate through the intestine
of Eisenia spp; (ii) the specific microbial changes dependent on
three different habitats (forest soil, compost, and vermiculture). To
achieve these aims, microbial characteristics were determined in
substrates, and in earthworm guts and faeces. The concomitant
earthworm microbiota was assessed using a combination of bio-
markers, molecular and culturing methods, to obtain information
concerning the total microbial biomass, community structure,
bacterial and archaeal composition, culturable bacteria-counts,
-growth strategy, and -species richness.
2. Material and methods
2.1. Earthworm and sample collection
The study is based on two to three independent observations for
each Eisenia population over three years. E.fetida earthworms were
collected from moist soil of a mixed forest brook rich in leaf litter
(mixed forest, Hlubok
a nad Vltavou, Czech Republic) in 2007, 2008
and 2009. E.andrei specimens were collected from a compost pile
with plant remains (
Cesk
eBud

ejovice, Czech Republic) in 2007 and
2009; the earthworms invaded the compost spontaneously from a
neighbour compost pile. The specimens of a second population of E.
andrei were collected from a large-scale vermiculture plant con-
sisting of straw, cattle manure and other organic agricultural wastes
(Mikul
cice, Czech Republic) in 2007 and 2008; the substrate was
inoculated using commercially acquired animals. Earthworm species
were identified on the base of species-specific mitochondrial gene
for cytochrome c oxidase subunit I [16]. Three composite soil sam-
ples, each consisting of three randomly collected partial samples,
were taken from a bottom of the brook in a depth 010 cm. Similarly,
three composite samples, each of three partial samples, were
collected from compost pile/vermiculture bed in a depth 020 cm
andstoredat4

C. Microbial characteristics of substrates (SUB) were
compared with those in the gut content (GT) and faeces (FS) of
earthworms. Six earthworms per one sample considered for gut
content analysis were used and the samples were acquired according
to Pi
zl and Nov
akov
a[33]. To obtain one sample of fresh faeces, nine
to twelve worms were placed on moist sterile filter paper over the
night and fresh excrement was collected into a sterile tube. Samples
were immediately performed to the chemical analyses or stored in
microtubes at minus 18

C for molecular analyses.
2.2. Soil, compost and vermiculture properties
Basic chemical and biochemical characteristics of soil (SO),
compost (CO), and vermiculture (VE) are summarized in Table 1.
The SO differed significantly in most of properties in comparison to
CO and VE. The soil pH was weakly acidic in contrast to neutral pH
Table 1
Chemical and biochemical parameters of forest soil, compost, and vermiculture substrates. Values are mean ±standard error. Chemical parameters were assessed in three
replicates (n ¼3); carbon- and humic-substances and enzymatic activity were analysed in two replicates (n ¼2).
Parameters Substrates
Forest soil Compost Vermiculture
pH (CaCl
2
) 5.7 ±0.1 7.4 ±0.1 6.9 ±0.1
Cox (%) 2.1 ±0.4 34.5 ±6.9 65.8 ±13.8
Ntot (%) 0.2 ±0.0 2.6 ±0.5 2.6 ±0.5
C/N 9.7 13.5 25.0
P(gkg
1
dw) 0.06 ±0.01 2.96 ±0.59 4.37 ±0.87
K(gkg
1
dw) 0.06 ±0.01 7.11 ±1.42 17.58 ±3.52
Mg (g kg
1
dw) 0.22 ±0.03 1.90 ±0.29 4.69 ±0.70
Ca (g kg
1
dw) 1.86 ±0.37 9.36 ±1.87 9.40 ±1.88
Water-soluble carbon (WSC) (%C) 0.08 ±0.06 a 0.33 ±0.14 a 0.52 ±0.17 a
Water-soluble carbohydrates (%C) 0.00 a 0.02 ±0.00 b 0.04 ±0.00 c
Extractable C
tot
(%C) 1.50 ±0.26 a 4.06 ±1.33 a 4.33 ±0.58 a
Humic acids (%C) 1.15 ±0.27 a 3.56 ±1.33 a 3.37 ±0.59 a
Fulvic acids (%C) 0.35 ±0.03 a 0.50 ±0.05 ab 0.96 ±0.15 b
Enzymatic activity
b
-glucosidase (
m
g p-nitrofenol g
1
h
1
) 165.7 ±39.5 a 167.8 ±1.2 a 284.1 ±1.9 a
Protease (
m
g consumed tyrosine g
1
h
1
) 43.1 ±3.6 a 67.8 ±1.4 b 100.7 ±0.2 c
Urease (mg NeNH
4
þ
g
1
h
1
) 0.95 ±0.12 a 4.58 ±0.10 b 4.66 ±0.85 b
Means in a row followed by variable lowercase letters are significantly different at P 0.05. Differences between means were evaluated by one-way ANOVA followed by
Tukey's post hoc test.
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e55 43

in CO and VE. The content of nitrogen (N
tot
) and other nutrient
elements were lower in SO than in CO/VE about one or even two
orders, respectively. Similarly there were measured lower organic
carbon (C
org
), water soluble carbohydrates and fulvic acids in SO
compared to CO/VE substrates. Nutrient and carbon poor soil
habitat was also characterized by lower enzymatic activity, espe-
cially urease and protease. The VE was in most of parameters (C
org
,
P, K, Mg, protease) the richest habitat characterized by more than
twice higher C/N ratio compared to CO and SO.
The pH was determined in 1:5 (v:v) soil/extract (0.01 M CaCl
2
extraction solution; [34]). Organic carbon was determined by dry
combustion and total nitrogen was determined by Kjeldahl digestion
[35]. Trace elements P, K, Mg, and Ca were analysed using flame
atomic absorption spectroscopy (FAAS). Water-soluble and extract-
able carbon and humic substances were analysed according to
modified methods described by Sanch
ez-Monedero et al. [36].Pro-
tease activity was measured by determining tyrosine consumed after
incubatingthe samples (1 g dw) with sodiumcaseinate (2%) for 2 h at
50

C. Optical density was measured at 700 nm after Folin-Ciocalteu
addition as a reagent to develop colour [37].Beta-glucosidaseac-
tivity was determined as p-nitrofenol released after incubating the
samples with p-nitrofenil-
b
-glucoside for 1 h at 37

C. Optical den-
sity was measured at 400 nm after developing the colour with 0.5 M
CaCl
2
[38]. Urease activity was assessed by measuring the NeNH
4
þ
content after incubating the samples with urea solution (0.2 M) for
2hat37

C and ammoniumextraction with 2 M KCl and 0.015 M HCl
in a spectrophotometer at 660 nm [39].
2.3. Fatty acid analysis
A simple PLFA analysis was used to quantify total microbial
biomass and microbial community composition [40] in all samples
during three years-observation. Simple PLFA analysis evaluates
ester-linked (EL-) PLFA without later fractionation of methylesters.
An extended PLFA method includes EL-PLFA indicative of aerobes,
and non-ester-linked (NEL-) PLFA interpreted as anaerobes. NEL-
PLFA group represents only minor part of soil microbiome PLFA
profile (e.g., 21% of total PLFA in fertilised soil [41]). Extended PLFA
procedure [42] was used for complex comparison of all samples in a
single experiment (2008). The extended PLFA analysis was repeated
in all sample types of E. fetida also in year 2009 to verify high
concentration of NEL-PLFA in faeces.
Fatty acid abbreviations (used in Fig. 1) according to Zelles [41]
denote the total number of carbon atoms followed by the number
of double bonds and their position from the aliphatic end (
u
). The
prefixes i- and a-refer to the terminal iso- and anteiso-branched
fatty acid chain, respectively. A number followed by Me-indicates
the position of the methyl group from the carboxyl end of the
molecule (i.e. mid-branching), and the prefix cy-represents cyclo-
propyl branching. The number preceding OH refers to the position
of the hydroxyl group of a hydroxy-fatty acid, and DMA refers to
aldehyde dimethylacetals. The individual PLFAs were classified
according to their composition and specific predominance in mi-
crobial taxa into groups (Table 2).
2.4. PCR-DGGE analysis of the archaeal and bacterial community
structure
DNA was extracted from 0.12 g earthworm gut content and
faeces and 0.5 g soil, compost and vermiculture substrates ac-
cording to Griffiths et al. [56]. The sample was placed in a Bio-101
Multimix 2 Matrix Tube (Qbiogene, Germany) and 0.5 ml CTAB
extraction buffer and 0.5 ml phenol-chloroform-isoamylalcohol
(25:24:1, v:v:v) were added. After the homogenisation step (40 s,
16,000 g) and centrifugation (5 min, 16,000 g), the aqueous phase
was transferred and phenol was removed by mixing with an equal
volume of chloroform-isoamylalcohol (24:1, v:v). The centrifuga-
tion step was repeated and total nucleic acid was extracted using
two volumes of 30% polyethylene glycol-1.6 M NaCl on ice for 2 h.
Subsequently, the centrifugation step was repeated (10 min) and
the supernatant was removed, DNA was air-dried and washed with
0.6 ml ice-cold 70% ethanol (centrifugation for 10 min) and again
air-dried. Extracted DNA was quantified with a spectrophotometer
(Genesys 6, Thermo Spectronic, USA) and subjected to electro-
phoresis in a 1% (w:v) agarose gel using 1 TAE buffer and
ethidium bromide staining [57].
PCR amplification of the target DNA was performed in a PCR
thermocycler (T 3000 Thermocycler, Biometra, Germany) using
different bacterial and archaeal primer sets. Each PCR mixture
contained 50 ng extracted DNA, 0.2
m
M each primer, 0.625 U Dream
Taq DNA Polymerase, 1 DNA polymerase buffer, 5
m
g Bovine
serum albumin (BSA), 0.2 mM dNTP-Mix and 3.5 mM MgCl
2
in a
final volume of 50
m
l (all MBI Fermentas, Lithuania).
Bacterial 16S rRNA genes were amplified using the universal 16S
Fig. 1. Relative distribution of phospholipid fatty acid (PLFA) functional groups in the PLFA profile evaluated in substrates (SUB), gut contents (GT), and faeces (FS) of three different
earthworm populations. The individual PLFAs were classified according to their source and structure into fatty acid groups: ester-linked (EL-) saturated (EL-STFA), monounsaturated
(EL-MUFA), polyunsaturated (EL-PUFA), terminally-branched (EL-t-Br-FA), mid-branched (EL-m-Br-FA), cyclopropyl-branched (EL-cy-FA), hydroxy-substituted (EL-OH-FA), and
branched hydroxy-substituted (EL-Br-OH-FA) fatty acids. The non-ester-linked fatty acids (NEL-PLFA) included the group of unsubstituted (NEL-UNSFA), hydroxy-substituted fatty
acids (NEL-OH-FA), and aldehyde dimethylacetals (DMA).
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e5544

rRNA primer set 984FGC (5
0
- [gc] GCACGGGGGGAACGCGAA-
GAACCTTAC e3
0
) and 1378R (5
0
- CGGTGTGTACAAGGCCCGGGAACG
e3
0
)[58]. The PCR included an initial 4 min denaturation at 95

C
and was followed by 10 thermal cycles of 1 min at 94

C, 1 min at
60

C and 2 min at 72

C followed by 25 thermal cycles of 1 min at
94

C, 1 min at 55

C and 2 min at 72

C. Amplification was
completed with a final extension step at 72

C for 15 min.
The archaea 16S rRNA genes were amplified according to Coolen
et al. [59] using primers Parch519F (5
0
- CAGCMGCCGCGGTAA e3
0
)
[60] and Arch915RGC (5
0
- [gc] GTGCTCCCCCGCCAATTCCT e3
0
)[61].
The PCR started with an initial denaturation step at 96

C for 5 min,
followed by 35 thermal cycles of 30 s at 94

C, 40 s at 57

C, and 40 s
at 72

C and a final extension step at 72

C for 10 min. PCR products
were visualised by electrophoresis in 1% (w:v) agarose gels and
ethidium bromide staining (10 mg ml
1
).
The DGGE was performed with the Ingeny PhorU2 system
(Ingeny International BV, The Netherlands) according to
Chro
n
akov
a et al. [62] using some modifications: for bacterial as
well as archaeal DGGE, 30
m
l PCR product was loaded onto 6% (w:v)
polyacrylamide gels with a denaturing gradient of 45%e60% and
45%e75% for bacteria and archaea, respectively, and was run at
100 V at 60

Cin1TAE-buffer (pH 7.4) for 16 h. After electro-
phoresis, the gels were stained with SYBR Green. A comprehensive
comparison of all samples was performed in a single experiment,
building on the preliminary experiment; the partial results are
summarized in methodological work by Koubov
a et al. [63].
2.5. Counts and identification of culturable bacteria
The dilution plate cultivation technique was used to estimate
the number of colony forming units (CFU). The development of CFU
during a one-week cultivation period on BBL Tripticase™Soy Broth
Agar medium (Becton Dickinson, USA) at 28

C was expressed as a
colony forming curve and used for a bacterial growth strategy
evaluation [64]. The identification of culturable bacteria was per-
formed by the MIS Sherlock System (TSBA 6, MIDI Inc., USA) after
previous preparation of the samples according to the manufac-
turer's protocol.
2.6. Statistical analysis
The analysis of variance (one-way ANOVA) and Tukey's post-hoc
test were used to detect the differences between the abundance
values of microbial biomass at P <0.05 within samples in conse-
quence of the gut passage in a single earthworm population.
KruskaleWallis test (P <0.05) and multiple comparisons of mean
ranks for all groups were used to find the significant differences
between CFU numbers since data were not normally distributed
(STATISTICA 7). The differences between important microbial
taxonomic groups were evaluated based on the relative abundance
of specific biomarkers or fatty acid groups within the PLFA profile
using one-way ANOVA and Tukey's test. Principal component
analysis (PCA) was applied to visualise the sample distribution
within the PLFA profile, log [mol% þ1] values were used (Canoco
for Windows 4.5, Centre for Biometry Wageningen, The
Netherlands). The ordination plot was created with CanoDraw for
Windows 4.5 [65]. One-way ANOVA and Tukey's post-hoc test of PC
sample scores were used to depict the effect of passage and the
earthworm population on the FA distribution.
DGGE banding patterns were normalised and analysed using the
GelCompar II software package, version 4.0 (Applied Maths, Ghent,
Belgium). Calculation of the pair-wise similarities was based on the
Dice correlation coefficient and an unweighed pair-group method
using arithmetic averages (UPGMA). The PCA of DGGE patterns was
used to explain the variability of the samples, and PCA sample
scores were tested using one-way ANOVA (P <0.05) and Tukey's
post-hoc test to reveal the differences between DGGE banding
patterns.
3. Results
3.1. Microbial biomass and PLFA microbial community profile
Viable microbial biomass, examined as EL-PLFA by simple PLFA
analysis, increased significantly by about one-to two-fold in
earthworm gut compared to substrates for all studied habitats
(Table 3,Table A.1). The EL-PLFA measured in faeces had an inter-
mediate response compared to that in substrates and guts; the
Table 2
Phosholipid fatty acid (PLFA) functional groups used in this study and their interpretation. The abbreviations of FA groups were used according to Zelles [41].
Abbreviation Indicator Interpretation Reference
EL-PLFA ester-linked fatty acid aerobic microorganisms [41,43]
NEL-PLFA non-ester-linked fatty acids anaerobic and extremophilic microorganisms [41,43]
EL-STFA saturated EL-PLFA all microorganisms [44]
EL-MUFA monounsaturated EL-PLFA aerobic eukaryotes and prokaryotes [45]
EL-PUFA polyunsaturated EL-PLFA microeukaryotes, rare in bacteria [44,46,47]
EL-t-Br-FA terminally branched EL-PLFA Gram-positive bacteria [48,49]
EL-m-Br-FA mid-branched saturated EL-PLFA actinomycetes [50,51]
EL-cy-FA cyclopropyl-branched saturated EL-PLFA Gram-negative bacteria [41,43]
EL-OH-FA hydroxy-substituted saturated EL-PLFA
with straight chain
Gram-negative bacteria, fungi [45,49]
EL-Br-OH-FA hydroxy-substituted saturated EL-PLFA
with branched chain
Gram-negative bacteria, mainly Bacteroidetes [45]
NEL-UNSFA unsubstituted NEL-PLFA anaerobes [45,52]
NEL-OH-FA hydroxy-substituted NEL-PLFA facultative anaerobes (Sphingomonas, Bacteroides, Flavobacterium,Candida)[45,5254]
DMA aldehyde dimethylacetals plasmalogen-containing strict anaerobes [52,55]
Table 3
Viable microbial biomass (EL-PLFA) evaluated in substrates (SUB), gut contents (GT),
and faeces (FS) of three different Eisenia populations: soil (SO), compost (CO) and
vermiculture (VE); (mean ±standard error; n ¼6).
EL-PLFA
[nmol PLFA g
1
dw]
SO CO VE
SUB 29.6 ±11.8 aA 73.1 ±43.7 aA 48.5 ±15.6 aA
GT 1216.0 ±286.8 bA 1335.3 ±267.9 bA 2095.9 ±426.0 bA
FS 1148.7 ±578.2 abA 656.3 ±95.8 aA 787.0 ±71.4 aA
Means followed by variable lowercase letters in a column are significantly different
at P 0.05; variable uppercase letters in a row show significantly different PLFA
mean values within earthworms. Differences between means were evaluated by
One-way ANOVA followed by Tukey's post hoc test.
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e55 45

values were not significantly different. The extended PLFA results
confirmed above described trend of quantitative microbial differ-
ences caused by passage of the substrates (Table A.2) and in more
details explained qualitative composition (Fig. 1,Fig. A.1). It showed
that substrates of all habitats had well-balanced proportions of
individual PLFA functional groups. Passage significantly affected
several groups. (i) The total NEL-PLFA relative abundance signifi-
cantly increased and accounted for 73%, 53%, and 31% of the total
microbial profile in faeces of soil, compost, and vermiculture
earthworms, respectively. In detail, the DMA (strict anaerobes)
relative abundance increased significantly in gut and faeces of soil
earthworm at the expense of NEL-OH-FA (facultative anaerobes).
(ii) The EL-PUFA accounted for a relatively low molar percentage in
soil and in compost substrate (4% and 6%, respectively) but
increased significantly in gut of soil and compost earthworms (to
21% and 13%, respectively) (Fig. 1). The EL-PUFA represented 13% in
all vermiculture samples. (iii) In contrast, the individual EL-PLFA
groups (MUFA, t-Br-FA, cy-FA), that accounted for a relatively
high molar percentage of the PLFA profile in substrates, were
suppressed in gut contents and faeces of all earthworm pop-
ulations. (iv) EL-OH-FA and EL-Br-OH-FA increased in gut and
faeces after the passage of vermiculture (Fig. 1). The multivariate
comparison (PCA) performed with the PLFA community profiles
extracted two axes explaining 80.6% of the total variance (Fig. A.1).
All the substrates were discriminated in a cluster around the
negative pole of the PC1 axis which explained 64.5% of the total
variance. All the GT samples and CO-FS samples were distributed
along the negative pole of the PC2 axis which explained 16.1% of the
variance. These samples were characterised by anaerobic markers
(NEL-UNSFA, 30e55% of the PLFA profile), by the fungal marker
18:2
u
6,9 (Fig. A.1), and evenly balanced groups of EL-t-Br-FA, EL-
PUFA, EL-MUFA, and EL-STFA (Fig. 1). Soil-earthworm-faeces sam-
ples (SO-FS) created one separate cluster, where NEL-UNSFAs and
DMAs extremely enriched the PLFA profile (Fig. 1,Fig. A.1). The PC1
axis separated the faeces of the vermiculture population (VE-FS)
from that of both soil and compost populations. The VE-FS samples
were clustered together with all samples of substrate. The PLFA
profile of VE-FS was well-balanced and the relative content of EL-
PLFA was significantly higher compared to the other FS samples
(Fig. 1). One-way ANOVA of the PC sample scores demonstrated a
significant effect (P <0.05) of passage on the PLFA microbial
community profile in all earthworm populations. The post-hoc test
showed a different microbial PLFA composition of FS samples
compared to both SUB and GT samples, whereas no differences
were found between SUB and GT samples in all three populations.
3.2. Archaeal and bacterial community structure
Cluster analysis of archaeal DGGE patterns revealed that passage
shifted the archaeal community structure in GT and FS compared to
Fig. 2. Dendrogram of archaeal DGGE fingerprints based on 16S rDNA extracted from substrates (SUB), gut content (GT) and faeces (FS) of forest (SO), compost (CO), and ver-
miculture (VE) earthworm populations. Profiles were clustered by Dice correlation and an unweighted pair-group method using arithmetic averages (UPGMA).
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e5546

SUB in all three earthworm populations (Fig. 2). This analysis of
fingerprints also depicted the lowest similarity among substrate,
gut, and faeces primarily in E.fetida inhabiting forest soil. PCA of
DGGE patterns extracted two PC axes explaining 35.0% of the total
variability (Fig. A.2a). One-way ANOVA of PC1 sample scores
confirmed significant differences (P <0.001) among SUB, GT, and FS
archaeal DGGE profiles of all Eisenia populations.
For bacterial banding patterns, passage showed a lower influ-
ence on the bacterial community structure (Fig. 3). Bacterial
community of soil (SO-SUB) was significantly different from that in
GT and FS samples. Compost and vermiculture substrate commu-
nities were more similar to those of GTand FS samples than those in
the archaeal fingerprint. Principal component analysis of DGGE
patterns extracted two PC axes explaining 36.0% of the total vari-
ability (Fig. A.2b). No evident separation of the samples was created
along the first two PC axes. One-way ANOVA of PC1 sample scores
revealed differences among SUB, GT, and FS bacterial DGGE banding
patterns of the soil population (P <0.001). A significant effect of
passage on the bacterial community was also found for the
compost population (P <0.01), although banding patterns were
only different in FS samples. Gut passage through the vermiculture
population had no significant impact on the bacterial DGGE
fingerprint. The effect of passage on bacterial composition
decreased along the gradient in the direction from nutrient-poor
forest soil to the richest vermiculture habitat.
3.3. Culturable bacteria-counts, growth strategy, and species
richness
The average counts of culturable bacteria were two to three
orders higher in the GT and FS of compost and vermiculture
earthworms compared to the GT and FS of soil earthworms and
three to four orders higher in compost and vermiculture substrates
Fig. 3. Dendrogram of bacterial DGGE fingerprints based on 16S rDNA extracted from substrates (SUB), gut content (GT) and faeces (FS) of forest (SO), compost (CO), and ver-
miculture (VE) earthworm populations. Profiles were clustered by Dice correlation and an unweighted pair-group method using arithmetic averages (UPGMA).
Table 4
Colony forming units (CFU) of cultivable bacteria in substrates (SUB), gut contents
(GT), and faeces (FS) of three different Eisenia populations: soil (SO), compost (CO)
and vermiculture (VE); (mean ±standard error; n ¼6).
Counts of cultivable bacteria
[CFU g
1
dw]
SO CO VE
SUB 5.6 ±1.2 10
6
aA 2.7 ±0.8 10
10
aB 5.8 ±0.8 10
9
aB
GT 2.8 ±0.1 10
7
bA 5.2 ±0.6 10
9
aB 1.0 ±0.3 10
10
abB
FS 1.7 ±0.1 10
7
abA 3.7 ±0.5 10
9
aB 1.2 ±0.1 10
10
bC
Means followed by variable lowercase letters in a column are significantly different
at P 0.05; variable uppercase letters in a row show significantly different CFU
mean values within earthworms. Differences between means were evaluated by
KruskaleWallis test followed by multiple comparisons of mean ranks for all groups.
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e55 47

compared to soil substrate. Earthworm passage significantly
increased the counts of culturable bacteria in the GT of the soil
earthworm population and in the FS of the vermiculture popula-
tion, no differences were found after compost passage (Table 4).
Soil earthworm also accelerated the bacterial growth rate in passed
material (Fig. 4a) in contrast to compost and vermiculture earth-
worms (Fig. 4b; Fig. 4c).
The richness of culturable bacterial species decreased during
passage in all earthworm populations (Fig. 5,Table A.3,Table A.4).
Bacterial species identification revealed the following differences
between samples derived from the three earthworm populations:
whereas soil compost and vermiculture substrates favoured by
Actinobacteria (e.g., Arthrobacter globiformis,Microbacterium sp.,
Nesterenkonia halobia) and Firmicutes (e.g., Bacillus sp., Paeniba-
cillus sp.), in the gut content and faeces of soil earthworms domi-
nated Gammaproteobacteria (e.g., Aeromonas sp., Pseudomonas sp.,
Salmonella typhimurium). Gut content of compost population
inhabited almost Firmicutes (e.g., Bacillus sp., Paenibacillus sp.) and
in faeces dominated Actinobacteria (e.g., Microbacterium sp.,
Micrococcus luteus,Oerskovia turbata,Rhodococcus rhodochrous).
Whereas, Gammaproteobacteria dominated in gut content of ver-
miculture earthworm, Firmicutes increased in faeces.
4. Discussion
4.1. Different feeding habitats
Various feeding habitats resulted in different passage effects of
indigenous soil earthworm and related compost and vermiculture
earthworms. Earthworm gut passage had a higher impact on the
microbial community of forest soil than on that of compost and
vermiculture. The difference in chemical and biochemical parame-
ters, especially in the organic C, water soluble carbohydrates and soil
organic matter appears as an important factor in various earthworm
responses in different habitats [66]. Distinct NEL-PLFA concentra-
tions in faeces of three earthworm populations could be explained
by different antimicrobial defense mechanisms among earthworm
species from various habitats [16]. Compost and vermiculture
earthworms are evolutionary adapted to substantially higher quan-
tity of microbes and their PLFA profile was well balanced between
gut and faeces. In the other hand, the intestine of soil E.fetida
selected and activated anaerobes that predominated in faeces.
The lower numbers of cultivable bacteria (CFU) in soil popula-
tion corresponded to the nutrient demands in forest soil habitat.
High amount of organic carbon in compost and vermiculture forms
these substrates more suitable for growth of microorganisms. Our
estimates of CFU in substrates were in agreement with those of
previous measurement of Dvo
r
ak et al. [16] where culturable bac-
terial numbers in soil and compost inhabited by Eisenia worms
reached 10
5
and 10
8
CFU g
1
dry matter, respectively. According to
previous studies, the counts of bacteria isolated from the gut in
compost earthworm E.fetida ranged from 10
6
to 10
9
CFU g
1
dry
matter [14,67,68], whereas the counts of culturable bacteria in the
Fig. 4. Growth strategy of culturable bacteria isolated from substrates (SUB), gut
contents (GT) and faeces (FS) of soil (a), compost (b), and vermiculture (c) earthworm
populations.
Fig. 5. Species richness of culturable bacteria and distribution in taxonomic groups.
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e5548

gut contents of other earthworm species from different ecological
groups (anecic Lumbricus terrestris, epigeic L.rubellus, and endogeic
Aporrectodea caliginosa and Octolasion lacteum), reached only
10
5
e10
6
CFU g
1
dry matter [9,10]. The acceleration of the bacterial
growth rate derived from gut and faeces of E.fetida correlates with
the relatively low CFU observed in forest soil. It is clear that the
nutrient-rich vermiculture substrate contained a large community
of fast-growing bacteria, a strategy that was preferred after
entering into an intestinal environment (Fig. 4). The richer the
habitat in nutrients and organic carbon was, the lower the influence
of earthworms to stimulate intestinal bacterial growth.
4.2. Passage effects
Previous studies showed that earthworm gut increases or does
not change the abundance of microbial biomass [68]. Our results
demonstrate that passage increased total microbial biomass as
indicated by a one-to two-fold increase in the EL-PLFA molar con-
centration in the gut and faeces of three Eisenia populations. In
accordance with our study, Sampedro and Whalen [7] found about
300 times greater total PLFA concentration in the gut of L.terrestris
than in bulk soil. Similarly, they found significant changes in
microbial-derived PLFA profiles of soil and gut and described that
the passage significantly increased the concentration of biomarkers
indicative for aerobic bacteria, microeukaryotes and fungi. The
passed material through the earthworm gut may differ between
species and this difference might result in various increments of gut
microbial biomass. For instance, some species such anecic L.ter-
restris feeds mainly upon intact organic wastes for nutrition,
whereas other species such epigeic E.fetida prefer organic matter in
an advanced stage of decomposition. Endogeic species e. g. Allolo-
bofora caliginosa even extract the nutrients from finely fragmented
organic matter mixed with soil [69]. Moreover, we showed the
significant increase of NEL-PLFA concentration in earthworm gut
and faeces. The increased concentration of the NEL-UNSFA group in
earthworm gut and faeces is mainly attributable to the presence of
anaerobes [52]. Compared to soil, the intestinal tract is a micro-
environment with a higher carbon, nitrogen and water content and
distinct oxygen deficiency [70]. Many anaerobic or facultatively
anaerobic bacteria (e.g., Clostridium, Aeromonas,Bacillus,Shewa-
nella, Propionibacterium, Staphylococcus, Paenibacillus and Photo-
bacterium) as well as archaea, are abundant in the guts of
earthworms [13,14,7173]. The conditions of the gut are suitable
for the activation of dormant or inactive microbial forms that might
be present in soil [70]. In this study, we obtained these results by
culturable bacteria counts and identification. We revealed that the
gut environment, mainly of compost earthworms, activated spore-
forming bacteria (Bacillus licheniformis, Bacillus megaterium,Bacillus
mycoides, Bacillus pumilus, Bacillus subtilis, Paneibacillus macerans,
etc.). The same findings were described previously by Fischer et al.
[74], who demonstrated that passage by L.terrestris enhanced the
germination of B.megaterium spores. It is evident for soil and ver-
miculture earthworm populations that the richness of the Gam-
maproteobacteria increased in the gut contents. The Gram-negative
facultative anaerobic bacteria, Aeromonas, together with Pseudo-
monas, which are able to grow in anaerobic conditions, have been
previously identified by culture-dependent and independent ap-
proaches as typical genera in the intestine of E.fetida [12,13].So
called ‘sleeping beauty paradox’, the contradiction between the
short generation time of microorganisms and their slow turnover
[75], underlines the fact that earthworm gut passage might stim-
ulate dormant microbiota.
Although culturable bacterial phyla differed in vermiculture gut
contents and faeces, the PCR-DGGE analysis did not confirmed the
differences in bacterial molecular fingerprints. Furlong et al. [19]
revealed in L.rubellus, that soil and cast bacterial isolates were
different but represented the same phylogenetic groups. It should
be taken into account that facultative anaerobes were found to be
as abundant in an anaerobic gut environment and these were not
identified by cultivation [71], thus the differences between cultur-
able aerobic heterotrophic bacterial communities were not obvious
in the resulting bacterial molecular profiles. Bacteria were distin-
guished by a higher diversity compared to archaea, and the differ-
ences that we observed might correspond to the changes in
dominant representatives, therefore, the total bacterial community
might have reacted less sensitively to environmental changes than
archaea, as was reported in previous study by Koubov
a et al. [63].
The archaeal DGGE patterns were strictly divergent in different
stages of passage. Furlong et al. [19] found that RFLP patterns
analysed in L.rubellus were different between soil and casts
archaeal clones. The differences in archaeal profiles might be
influenced by typical soil and compost methanogenic and non-
methanogenic Euryarchaeota as well as NH
4
-oxidising Thau-
marchaeota [76]. Nevertheless, many uncultured representatives of
these two groups exist, whose functional properties are unknown.
Koubov
a et al. [42] showed an earthworm-mediated decrease in
Methanosarcina sp. abundance and methanogenic diversity
changes in soil altered by E.andrei; nevertheless, the quantity of
total methanogens was stable. A recent study by Depkat-Jakob et al.
[18] demonstrated that methanogenic taxa Methanosarcinaceae,
Methanobacteriaceae, and Methanomicrobiaceae might be associ-
ated with the emission of methane by Eudrilus eugeniae. Meth-
anogenic and ammonium-oxidizing archaea might be the
dominant archaeal groups attending composting processes [77],
however, the participation of individual archaeal taxa on earth-
worm ingestion processes remains unresolved. Microbial species
selection and reduction during earthworm gut passage could play a
key role in biological soil amendments like bioturbation and
vermicomposting.
5. Conclusions
In summary, we have shown that the passage of consumed
material through the gut of Eisenia spp. increased total viable mi-
crobial biomass, changed the microbial structure and reduced the
richness of the microbial communities in gut and faeces. We have
revealed that the influence of Eisenia spp. on associate microor-
ganisms depends on the earthworm feeding habitat. Passage
affected the intestinal microbiome of indigenous soil earthworm in
more aspects than related compost and vermiculture populations:
the results indicated greater changes in archaeal community
structure, higher abundance of anaerobes in passed forest soil. The
richer the habitat in nutrients and organic carbon was, the lower
the influence of earthworm gut passage on bacterial growth rate
has been found. This work brought original reveals about archaeal
community and total anaerobes accompanying intestinal tract of
Eisenia spp. Further molecular screening of the microbial commu-
nity might help the understanding of the microbial functions
associated with Eisenia spp. earthworms.
Acknowledgements
We are grateful to M. 
Silerov
a for the molecular identification of
earthworms. We thank N. Serrami
a for technical support in soil and
compost biochemical analyses. The study was supported by the
Grant Agency of the Academy of Sciences of the Czech Republic
(project No. IAA600200704) and by the Ministry of Education,
Youth, and Sports of the Czech Republic (project No. LC06066).
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e55 49

Appendices
Fig. A.1. Ordination plot of sample distribution after principal component analysis of the complex PLFA profile. Only PLFAs that fitted with PC1 and PC2 by more than 40% are
projected in the figure. The symbols represent: soil (SO), compost (CO), vermiculture (VE), substrate (SUB), earthworm gut contents (GT) and earthworm faeces (FS). The arrows
demonstrate EL- and NEL-PLFAs.
Fig. A.2. Principal component analysis of DGGE banding patterns for archaea (a), and bacteria (b). The symbols represent: soil (SO), compost (CO), vermiculture (VE), substrate
(SUB), earthworm gut contents (GT), and earthworm faeces (FS). Values on the axes indicate the percentage variability in the DGGE banding patterns.
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e5550

Table A.1
PLFA microbial functional groups [nmol PLFA g
1
dw] and substrate availability to microbes evaluated in substrates (SUB), gut contents (GT), and faeces (FS) of three different Eisenia populations: soil (SO), compost (CO) and
vermiculture (VE); (mean ±standard error; n ¼6). Specific PLFA biomarkers were used to estimate the aerobic bacteria (i15:0; a15:0; 15:0; 16:1
u
7c; i17:0; cy17:0; cy19:0), actinomycetes (10Me18:0; 10Me17:0; 10Me16:0), fungi
(18:2
u
6; 18:1
u
9), and aerobic microeukaryotes (20:4
u
6; 20:2
u
6; 18:3
u
6; 18:2
u
6). The substrate availability to microbes was estimated as the ratio EL-MUFA/EL-STFA.
Microbial parameters Samples analysed
SO CO VE
SUB GT FS SUB GT FS SUB GT FS
Aerobic
a
bacteria 12.7 ±5.3 a 226.4 ±82.2 a 149.6 ±100.2 a 25.0 ±16.1 a 296.3 ±86.5 b 69.1 ±38.7 a 25.0 ±8.8 a 460.2 ±92.1a 226.0 ±24.5 a
Actinomycetes
b
1.1 ±0.3a 23.2 ±20.5 a 54.8 ±38.0 a 4.1 ±2.6 a 24.5 ±15.6 a 14.9 ±9.7 a 0.8 ±0.3 a 0.0 ±0.0 a 0.3 ±0.3 a
Aerobic
c
microeukaryotes 2.3 ±1.2 a 180.3 ±91.5 a 252.8 ±127.1 a 4.7 ±2.8 a 181.3 ±120.8 a 43.9 ±27.9 a 7.1 ±3.5 a 614.5 ±162.3 b 93.1 ±19.3 a
Fungi
a
3.0 ±1.6 a 181.5 ±58.8 b 30.1 ±22.9 a 10.6 ±0.9 a 157.1 ±34.4 b 85.1 ±5.8 ab 4.1 ±2.0 a 232.3 ±61.7 b 44.8 ±5.3 a
Substrate
d
availability 0.8 ±0.3 a 0.4 ±0.2 a 0.7 ±0.2 a 0.6 ±0.2 a 0.3 ±0.2 a 0.5 ±0.3 a 1.1 ±0.2 a 0.5 ±0.0 b 0.7 ±0.2 ab
Means in a row followed by variable lowercase letters are significantly different at P 0.05 within samples in a single earthworm population. Differences within samples in consequence of the gut passage in a single earthworm
population were evaluated by one-way ANOVA followed by Tukey's post hoc test.
a
Frostegård and Bååth (1996) [78].
b
Kroppenstedt (1985) [50].
c
Erwin (1973) [79].
d
Bossio and Scow (1998) [80].
Table A.2
Abundance PLFA functional groups [nmol PLFA g
1
dw] in the PLFA profile evaluated in substrates (SUB), gut contents (GT), and faeces (FS) of three different Eisenia populations: soil (SO), compost (CO) and vermiculture (VE);
(mean ±standard error; n ¼2). Measurements were obtained from detailed extended PLFA analysis from observations over one year.
PLFA functional groups Samples analysed
SO CO VE
SUB GT FS SUB GT FS SUB GT FS
EL-STFA 2.9 ±0.3 a 193.5 ±46.3 b 87.0 ±16.6 ab 37.1 ±2.5 a 350.0 ±26.8 b 180.3 ±0.8 c 13.7 ±1.2 a 198.5 ±42.0 a 119.4 ±36.0 a
EL-MUFA 7.2 ±1.5 a 272.4 ±15.7 b 26.9 ±5.7 a 78.4 ±3.5 a 521.7 ±157.5 a 386.8 ±57.5 a 39.0 ±2.5 a 246.7 ±94.2 a 234.3 ±42.1 a
EL-PUFA 0.9 ±0.2 a 331.2 ±8.8 b 75.8 ±37.8 a 13.9 ±1.7 a 544.1 ±146.4 b 131.8 ±10.6 ab 12.9 ±1.8 a 193.6 ±63.7 a 116.7 ±36.5 a
EL-t-Br-FA 3.6 ±0.5 a 257.8 ±41.2 b 92.8 ±10.5 a 47.4 ±2.5 a 350.6 ±69.4 b 129.8 ±14.3 a 22.1 ±1.5 a 230.9 ±69.5 b 104.3 ±4.6 a
EL-m-Br-FA 0.7 ±0.1 a 8.4 ±6.4 a 2.5 ±2.5 a 12.4 ±0.3 a 73.5 ±5.4 b 44.6 ±9.0 ab 0.5 ±0.0 a 0.0 ±0.0 a 0.7 ±0.8 a
EL-cy-FA 0.7 ±0.1 a 6.0 ±4.3 a 0.0 ±0.0 a 16.7 ±0.6 a 17.5 ±3.0 a 11.4 ±1.7 a 1.8 ±0.2 a 2.5 ±0.2 a 3.4 ±2.1 a
EL-OH-FA 0.1 ±0.0 a 5.3 ±0.8 b 0.7 ±0.7 a 1.6 ±0.3 a 10.9 ±3.1 ab 15.1 ±1.2 b 0.1 ±0.1 a 50.3 ±18.2 a 25.8 ±9.5 a
EL-Br-OH-FA 0.0 ±0.0 a 1.7 ±0.2 b 1.6 ±0.4 b 0.7 ±0.0 a 28.4 ±0.0 b 12.7 ±1.5 c 0.3 ±0.0 a 23.6 ±2.4 b 11.2 ±0.3 c
EL-PLFA
tot
16.2 ±2.9 a 1076.3 ±124.8 b 287.4 ±61.6 a 210.6 ±11.3 a 1901.1 ±419.3 b 917.6 ±96.2 ab 91.0 ±7.2 a 946.0 ±163.6 b 616.7 ±128.1 ab
NEL-UNSFA 4.6 ±0.2 a 499.4 ±64.4 ab 736.6 ±140.7 b 26.1 ±0.5 a 1945.2 ±412.3 b 933.2 ±137.0 ab 11.0 ±2.5 a 471.1 ±69.8 b 268.6 ±77.6 ab
NEL-OH-FA 0.8 ±0.0 a 6.1 ±1.5 b 4.0 ±0.2 ab 7.9 ±0.2 a 213.8 ±100.3 a 76.0 ±13.7 a 3.6 ±0.7 a 61.0 ±26.6 a 5.8 ±0.2 a
DMA 0.2 ±0.0 a 38.5 ±3.0 b 50.9 ±27.1 b 0.0 ±0.0 a 0.0 ±0.0 a 0.0 ±0.0 a 2.0 ±0.5 a 14.0 ±5.2 a 6.5 ±6.5 a
NEL-PLFA
tot
5.6 ±0.2 a 543.9 ±65.8 ab 791.5 ±167.9 b 34.0 ±0.7 a 2159.0 ±512.6 b 1009.2 ±150.8 ab 16.6 ±2.7 a 546.1 ±101.6 b 280.9 ±83.9 ab
PLFA
tot
21.8 ±2.7 a 1620.2 ±190.6 b 1078.9 ±106.4 ab 244.6 ±10.6 a 4060.0 ±931.9 b 1926.7 ±246.9 ab 107.6 ±4.6 a 1492.1 ±265.2 b 897.6 ±212.0 ab
Means in a row followed by variable lowercase letters are significantly different at P 0.05 within samples in consequence of the gut passage in a single earthworm population. Differences between means were evaluated by
one-way ANOVA followed by Tukey's post hoc test.
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e55 51

Table A.3
Cultivable species of Actinobacteria and Firmicutes isolated from substrates (SUB), gut contents (GT), and faeces (FS) in three different Eisenia populations: soil (SO), compost
(CO) and vermiculture (VE).
Phyla and species Soil Compost Vermiculture
SUB GT FS SUB GT FS SUB GT FS
Actinobacteria
Arthrobacter aurescens þ
Arthrobacter globiformis þþþ
Arthrobacter oxydans þ
Brevibacterium lyticum þ
Cellulomonas fimi þ
Cellulomonas flavigena þ
Cellulomonas gelida þ
Cellulosimicrobium cellulans þþþ
Corynebacterium xerosis þ
Curtobacterium flaccumfaciens þþþ
Dactylosporangium fulvum þ
Kocuria rhizophila þ
Kocuria rosea þþþ
Kocuria varians þ
Lechevalieria flava þþ
Microbacterium barkeri þ þ
Microbacterium esteraromaticum þ
Microbacterium flavescens þ
Microbacterium chocolatum þþ
Microbacterium lacticum þ
Microbacterium laevaniformans þ
Microbacterium liquefaciens þ
Microbacterium trichothecenolyticum þ
Microbispora diastatica þ
Micrococcus luteus þþ
Micrococcus lylae þ
Nesterenkonia halobia þþþ
Nocardiopsis dassonvillei dassonvillei þ
Oerskovia turbata þþ
Rhodococcus erythropolis þþ
Rhodococcus fascians þ
Rhodococcus rhodochrous þþ
Streptomyces biverticillatus þ
Streptoverticilium reticulum þþ
Species richness 8 5 3 4 3 7 12 8 2
Firmicutes
Aneurinibacillus migulanus þþ
Bacillus atrophaeus þþþ
Bacillus cereus þþþ þ
Bacillus clausii þþ
Bacillus coagulans þ
Bacillus lentus þ
Bacillus licheniformis þþþ þ þ
Bacillus megaterium þþþ
Bacillus mycoides þþ þ
Bacillus pumilus þþþþþ
Bacillus sphaericus þþþ
Bacillus subtilis þþþþ
Bacillus thuringiensis kurstakii þ
Bacillus viscosus þ þ
Brevibacillus reuszeri þþ
Carnobacterium piscicola þ
Listeria innocua þ
Paenibacillus lautus þ
Paenibacillus macerans þþþþ
Paenibacillus polymyxa þþþþþþ
Staphylococcus cohnii cohnii þ
Staphylococcus sciuri þ
Staphylococcus simulans þ
Virgibacillus pantothenticus þ
Species richness 9 3 1 6 7 4 14 5 9
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a et al. / European Journal of Soil Biology 68 (2015) 42e5552

References
[1] K. Svensson, H. Friberg, Changes in active microbial biomass by earthworms
and grass amendments in agricultural soils, Biol. Fertil. Soils 44 (2007)
223e228.
[2] M. Aira, J. Domínguez, Earthworm effects without earthworms: inoculation of
raw organic matter with worm-worked substrates alters microbial commu-
nity functioning, Plos One 6 (2011) e16354.
[3] S. Scheu, Changes in microbial nutrient status during secondary succession
and its modification by earthworms, Oecologia 84 (1990) 351e358.
[4] M. Egert, S. Marhan, B. Wagner, S. Scheu, M.W. Friedrich, Molecular profiling
of 16S rRNA genes reveals diet-related differences of microbial communities
in soil, gut, and casts of Lumbricus terrestris L. (Oligochaeta: Lumbricidae),
FEMS Microbiol. Ecol. 48 (2004) 187e197.
[5] N.V. Khomyakov, S.A. Kharin, T.Yu. Nechitailo, P.N. Golyshin, A.V. Kurakov,
B.A. Byzov, D.G. Zvyagintsev, Reaction of microorganisms to the digestive fluid
of earthworms, Microbiology 76 (2007) 45e54.
[6] L. Sampedro, R. Jeannotte, J.K. Whalen, Trophic transfer of fatty acids from gut
microbiota to the earthworm Lumbricus terrestris L. Soil Biol. Biochem. 38
(2006) 2188e2198.
[7] L. Sampedro, J.K. Whalen, Changes in the fatty acid profiles through the
digestive tract of the earthworm Lumbricus terrestris L. Appl. Soil Ecol. 35
(2007) 226e236.
[8] S. Marhan, E. Kandeler, S. Scheu, Phospholipid fatty acid profiles and xylanase
activity in particle size fractions of forest soil and casts of Lumbricus terrestris
L. (Oligochaeta, Lumbricidae), Appl. Soil Ecol. 35 (2007) 412e422.
[9] G.R. Karsten, H.L. Drake, Comparative assessment of the aerobic and anaerobic
microfloras of earthworm guts and forest soils, Appl. Environ. Microbiol. 61
(1995) 1039e1044.
[10] V. Kri
st
ufek, K. Ravasz, V. Pi
zl, Changes in densities of bacteria and microfungi
during gut transit in Lumbricus rubelus and Aporrectodea caliginosa (Oli-
gochaeta: Lumbricidae), Soil Biol. Biochem. 24 (1992) 1499e1500.
[11] M. G
omez-Brand
on, M. Lores, J. Dominguez, Species-specific effects of epigeic
earthworms on microbial community structure during first stages of
decomposition of organic matter, Plos One 7 (2012) e31895.
[12] S.W. Hong, I.S. Kim, J.S. Lee, K.S. Chung, Culture-based and denaturing
gradient gel electrophoresis analysis of the bacterial community structure
from the intestinal tracts of earthworms (Eisenia fetida), J. Microbiol. Bio-
technol. 21 (2011) 885e892.
[13] S.W. Hong, J.S. Lee, K.S. Chung, Effect of enzyme producing microorganisms on
the biomass of epigeic earthworms (Eisenia fetida) in vermicompost, Bio-
resour. Technol. 102 (2011) 6344e6347.
[14] K. Toyota, M. Kimura, Microbial community indigenous to the earthworm
Table A.4
Cultivable species of Proteobacteria and Bacteroides isolated from substrates (SUB), Cultivable species of Proteobacteria and Bacteroides isolated from substrates (SUB), gut
contents (GT), and faeces (FS) in three different Eisenia populations: soil (SO), compost (CO) and vermiculture (VE).
Phyla and species Soil Compost Vermiculture
SUB GT FS SUB GT FS SUB GT FS
a
-Proteobacteria
Brevundimonas diminuta þ
Brevundimonas vesicularis þ
Hyphomonas hirschiana þ
Rhizobium radiobacter þþ
Sphingomonas paucimobilis þ
Species richness 0 0 0 2 0 0 1 2 1
b
-Proteobacteria
Aquaspirillum autotrophicum þ
Delftia acidovorans þ
Duganella zoogloeiodes þ
Hydrogenophaga pseudoflava þ
Species richness 0 1 0 2 0 0 1 0 0
g
-Proteobacteria
Aeromonas ichthiosmia/hydrophila þþ þ þ
Aeromonas jandaei þ
Aeromonas salmonicida achromogenes þþ
Aeromonas salmonicida-masoucida þþþ
Aeromonas veronii þ þ
Enterobacter intermedius þþ
Chryseomonas luteola þ
Lysobacter antibioticus þþþþ
Lysobacter sp. þ
Pantoea agglomerans (Enterobacter) þ
Photobacterium angustum þ
Pseudomonas agarici þ
Pseudomonas alcaligenes þ
Pseudomonas fluorescens/mandelii þþþ
Pseudomonas fluorescens/taetrolens þ
Pseudomonas chlororaphis þ
Pseudomonas putida þ
Pseudomonas putida/vancouverensis þþ
Pseudomonas syringae-syringae þ
Pseudomonas syringae-tabaci þ
Pseudomonas syringae-tomato þ
Pseudoxanthomonas broegbernensis þþ
Pseudoxanthomonas sp. þ
Raoultella terrigena þ
Salmonella typhimurium þþþ
Serratia plymuthica þ
Shewanella putrefaciens þ
Shewanella putrefaciens/algae þþ
Stenotrophomonas acidaminiphila þþ
Vibrio fischeri þ
Yersinia aldovae þ
Species richness 5 12 7 1 0 1 7 10 7
Bacteroides
Chryseobacterium balustinum þ
Species richness 1 0 0 0 0 0 0 0 0
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e55 53

Eisenia foetida, Biol. Fertil. Soils 31 (2000) 187e190.
[15] J. Domíngues, A. Velando, A. Ferreiro, Are Eisenia fetida (Savigny, 1826) and
Eisenia andrei Bouch
e (1972) (Oligochaeta, Lumbricidae) different biological
species? Pedobiologia 49 (2005) 81e87.
[16] J. Dvo
r
ak, V. Man
cíkov
a, V. Pi
zl, D. Elhottov
a, M. 
Silerov
a, R. Roubalov
a,
F. 
Skanta, P. Proch
azkov
a, M. Bilej, Microbial environment affects innate im-
munity in two closely related earthworm species Eisenia andrei and Eisenia
fetida, Plos One 8 (2013) e79257.
[17] M. P
erez-Losada, J. Eiroa, S. Mato, J. Domínguez, Phylogenetic species
delimination of the earthworms Eisenia fetida (Savigny, 1826) and Eisenia
andrei Bouch
e, 1972 (Oligochaeta, Lumbricidae) based on mitochondrial and
nuclear DNA sequences, Pedobiologia 49 (2005) 317e324.
[18] P.S. Depkat-Jakob, S. Hunger, K. Schulz, G.G. Brown, S.M. Tsai, H.L. Drake,
Emission of methane by Eudrilus eugeniae and other earthworms from Brazil,
Appl. Environ. Microbiol. 78 (2012) 3014e3019.
[19] M.A. Furlong, D.R. Singleton, D.C. Coleman, W.B. Whitman, Molecular and
culture-based analyses of prokaryotic communities from an agricultural soil
and the burrows and casts of the earthworm Lumbricus rubellus, Appl. Envi-
ron. Microbiol. 68 (2002) 1265e1279.
[20] D. Thakuira, O. Schmidt, D. Finan, D. Egan, F.M. Doohan, Gut wall bacteria of
earthworms: a natural selection process, ISME J. 4 (2010) 357e366.
[21] B. Byzov, T.Yu. Nechitaylo, B.K. Bumazhkin, A.V. Kurakov, P.N. Golyshin,
D.G. Zvyagintsev, Culturable microorganisms from the earthworm digestive
tract, Microbiology 78 (2009) 360e368.
[22] J.R. Albani, S. Demuynck, F. Grumiaux, A. Lepretre, Fluorescence fingerprints of
Eisenia fetida and Eisenia andrei, Photochem. Photobiol. 78 (2003) 599e602.
[23] V. Pi
zl, 
Zí
zaly 
Cesk
e republiky (Earthworms of the Czech Republic), Sborník
p
rírodov
edeck
eho klubu v Uhersk
em Hradi
sti, Supplementum 9 (2002)
1e154 (in Czech).
[24] A. Vivas, B. Moreno, S. GarciaRodriguez, E. Benites, Assessing the impact of
composting and vermicomposting on bacterial community size and structure,
and microbial functional diversity of an olive-mill waste, Bioresour. Technol.
100 (2009) 1319e1326.
[25] Z.A. Hickman, B.J. Reid, Earthworm assisted bioremediation of organic con-
taminants, Environ. Int. 34 (2008) 1072e1081.
[26] A.Y. Renoux, M. Sarrazin, J. Hawari, G.I. Sunahara, Transformation of
2,4,6trinitrotoluene in soil in the presence of the earthworm Eisenia andrei,
Environ. Toxicol. Chem. 19 (2000) 1473e1480.
[27] S.M. Contreras-Ramos, D. 
Alvarez-Bernal, L. Dendooven, Characteristics of
earthworms (Eisenia fetida) in PAHs contaminated soil amended with sewage
sludge or vermicompost, Appl. Soil Ecol. 41 (2009) 269e276.
[28] A. Velando, J. Eiroa, J. Dominguez, Brainless but not clueless: earthworms
boost their ejaculates when they detect fecund, non-virgin partners, Proc. R.
Soc. B 275 (2008) 1067e1072.
[29] A. Beschin, M. Bilej, F. Hanssens, J. Raymakers, E. van Dyck, H. Revets, L. Brys,
J. Gomez, P. de Baetselier, M. Timmermans, Identification and cloning of a
glucan- and liopoplysaccharide-binding protein from Eisenia foetida earth-
worm involved in the activation of prophenoloxidase cascade, J. Biol. Chem.
273 (1998) 24948e24954.
[30] M. Bilej, P. de Baetselier, A. Beschin, Antimicrobial defense of earthworm, Folia
Microbiol. 45 (2000) 283e300.
[31] M. Bilej, P. de Baetselier, E. van Dijck, B. Stijlemans, A. Colige, A. Beschin,
Distinct carbohydrate recognition domains of an invertebrate defense mole-
cule recognize gram-negative and gram-positive bacteria, J. Biol. Chem. 276
(2001) 45840e45847.
[32] P. Proch
azkov
a, M. 
Silerov
a, B. Stijlemans, M. Dieu, P. Halada, R. Joskov
a,
A. Beschin, P. de Baetselier, M. Bilej, Evidence for proteins involved in
prophenoloxidase cascade Eisenia fetida earthworms, J. Comp. Physiol. B 176
(2006) 581e587.
[33] V. Pi
zl, A. Nov
akov
a, Interactions between microfungi and Eisenia andrei
(Oligochaeta) during cattle manure vermicomposting, Pedobiologia 47 (2003)
895e899.
[34] J. Zbíral, I. Honsa, et al., Soil Analyses, Part I, Central Institute for Supervising
and Testing in Agriculture, Brno, 2010 (in Czech).
[35] J. Zbíral, I. Honsa, S. Malý, Soil Analyses, Part III, Central Institute for Super-
vising and Testing in Agriculture, Brno, 1997 (in Czech).
[36] M.A. Sanch
ez-Monedero, A. Roig, C. MartínezPardo, J. Cegarra, C. Paredes,
A microanalysis method for determining total organic carbon in extracts of
humic substances. Relationships between total organic carbon and oxidable
carbon,, Bioresour. Technol. 57 (1996) 291e295.
[37] J.N. Ladd, J.H.A. Butler, Short-term assays of soil proteolytic enzyme activities
using proteins and dipeptide derivatives as substrates, Soil Biol. Biochem. 4
(1972) 19e30.
[38] F. Eivazi, M.A. Tabatabai, Glucosidases and galactosidases in soils, Soil Biol.
Biochem. 20 (1988) 601e606.
[39] E. Kandeler, H. Gerber, Short-term assay of soil urease activity using colori-
metric determination of ammonium, Biol. Fertil. Soils 6 (1988) 68e72.
[40] D. Elhottov
a, V. Kri
st
ufek, S. Malý, J. Frouz, Rhizosphere effect of colonizer
plant species on the development of soil microbial community during primary
succession on postmining sites, Commun. Soil Sci. Plan. 40 (2009) 758e770.
[41] L. Zelles, Fatty acid patterns of phospholipids and lipopolysaccharides in the
characterisation of microbial communities in soil: a review, Biol. Fertil. Soils
29 (1999) 111e129.
[42] A. Koubov
a, M. Goberna, M. 
Simek, A. Chro
n
akov
a, V. Pi
zl, H. Insam,
D. Elhottov
a, Effects of the earthworm Eisenia andrei on methanogens in a
cattle-impacted soil: a microcosm study, Eur. J. Soil Biol. 48 (2012) 32e40.
[43] L. Zelles, Identification of single cultured micro-organisms based on their
whole-community fatty acid profiles, using an extended extraction procedure,
Chemosphere 39 (1999) 665e682.
[44] O. Oravecz, D. Elhottov
a, V. Kri
st
ufek, V. 
Sustr, J. Frouz, J. T
ríska, K. M
arialigeti,
Application of ARDRA and PLFA analysis in characterizing the bacterial com-
munities of the food, gut and excrement of saprophagous larvae of Penthetria
holosericea (Diptera: Bibionidae): a pilot study, Folia Microbiol. 49 (2004)
83e93.
[45] L. Zelles, Phospholipid fatty acid profiles in selected members of soil microbial
communities, Chemosphere 35 (1997) 275e294.
[46] T.W. Federle, Microbial distribution in soil - new techniques, in: F. Megusar,
M. Ganthar (Eds.), Perspectives in Microbial Ecology, Slovene Society for
Microbiology, 1986, pp. 493e498. Ljubljana.
[47] D.M. L€
osel, Fungal lipids, in: C. Ratledge, S.G. Wilkinson (Eds.), Microbial Lipids
vol. 1, Academic Press, London, 1988, pp. 699e806.
[48] W.M. O'Leary, S.G. Wilkinson, Gram-positive bacteria, in: C. Ratlege,
S.G. Wilkinson (Eds.), Microbial Lipids, Academic Press, London, 1988, pp.
117e201.
[49] M.A. Cavigelli, G.P. Robertson, M.J. Klug, Fatty acid methyl ester (FAME) pro-
files as measures of soil microbial structure, in: H.P. Collins, G.P. Robertson,
M.J. Klug (Eds.), The Significance and Regulation of Soil Biodiverzity, Kluwer
Academic Publisher, The Netherlands, 1995, pp. 99e113.
[50] R.M. Kroppenstedt, Fatty acids and menaquinone analysis of actinomycetes
and related organisms, in: M. Goodfellow, D.E. Minnikin (Eds.), Chemical
Methods in Bacterial Systematics, Academic Press, London, 1985, pp.
173e189.
[51] M.P. Lechevalier, C. Debievre, H.A. Lechevalier, Chemotaxonomy of aerobic
actinomycetes: phospholipid composition, Biochem. Syst. Ecol. 5 (1977)
249e260.
[52] B.-M. Wilke, A. Gattinger, E. Fr€
ohlich, L. Zelles, P. Gong, Phospholipid fatty acid
composition of a 2,4,6-trinitrotolune contaminated soil and an uncontami-
nated soil as affected by a humification remediation process, Soil Biol. Bio-
chem. 36 (2004) 725e729.
[53] G.B. Wells, R.C. Dickson, L.R. Lester, Isolation and composition of
inositolphosphorylceramide-type sphingolipids of hyphal forms of Candida
albicans, J. Bacteriol. 178 (1996) 6223e6226.
[54] D.L. Balkwill, G.R. Drake, R.H. Reeves, J.K. Fredrickson, D.C. White,
D.B. Ringelberg, D.P. Chandler, M.F. Romine, D.W. Kennedy, C.M. Spadoni,
Taxonomic study of aromatic-degrading bacteria from deep-terrestrial-
subsurface sediments and description of Sphingomonas aromaticivorans sp.
nov., Sphingomonas subterranea sp. nov., and Sphingomonas stygia sp. nov, Int.
J. Syst. Bactriol 47 (1997) 191e201.
[55] D.C. White, J.O. Stair, D.B. Ringelberg, Quantitative comparisons of in situ
microbial biodiversity by signature biomarker analysis, J. Ind. Microbiol. 17
(1996) 185e196.
[56] R.I. Griffiths, A.S. Whiteley, A.G. O'Donnell, M.J. Bailey, Rapid method for
coextraction of DNA and RNA from natural environments for analysis of ri-
bosomal DNA- and rRNA-based microbial community composition, Appl.
Environ. Microbiol. 66 (2000) 5488e5491.
[57] J. Sambrook, D.W. Russell, Molecular Cloning: A Laboratory Manual, third ed.,
Cold Spring Harbour Laboratory Press, New York, 2001.
[58] H. Heuer, M. Krsek, P. Baker, K. Smalla, E.M. Wellington, Analysis of actino-
mycete communities by specific amplification of genes encoding 16S rRNA
and gel-electrophoretic separation in denaturing gradients, Appl. Environ.
Microbiol. 63 (1997) 3233e3241.
[59] M.J.L. Coolen, E.C. Hopmans, W.I.C. Rijpstra, G. Muyzer, S. Schoulen,
J.K. Volkman, J.S.S. Damst
e, Evolution of the methane cycle in Ace Lake
(Antarctica) during the Holocene: response of methanogens and methano-
trophs to environmental change, Org. Geochem 35 (2004) 1151e1167.
[60] L. Øvreås, L. Forney, F.L. Daae, V. Torsvik, Distribution of bacterioplankton in
meromictic Lake Saelenvannet, as determined by denaturing gradient gel
electrophoresis of PCR-amplified gene fragments coding for 16S rRNA, Appl.
Environ. Microbiol. 63 (1997) 3367e3373.
[61] D.A. Stahl, R.I. Amann, Development and application of nucleic acid probes in
bacterial systematics, in: E. Stackenbrandt, M. Goodfellow (Eds.), Sequencing
and Hybridization Techniques in Bacterial Systematics, John Wiley, Chi-
chester, 1991, pp. 205e248.
[62] A. Chro
n
akov
a, J. Ascher, J. Jirout, M.T. Ceccherini, D. Elhottov
a,
G. Pietramellara, M. 
Simek, Cattle impact on composition of archaeal, bacte-
rial, and fungal communities by comparative fingerprinting of total and
extracellular DNA, Biol. Fertil. Soils 49 (2013) 351e361.
[63] A. Koubov
a, B.A. Knapp, H. Insam, V. Pi
zl, D. Elhottov
a, The effect of passage
through the gut of earthworms (Annelida: Lumbricidae) on the diversity of
archaea and bacteria,, Acta Soc. Zool. Bohem 74 (2010) 69e74.
[64] V. Kri
st
ufek, D. Elhottov
a, A. Chro
n
akov
a, I. Dost
alkov
a, T. Picek, J. Kal
cík,
Growth strategy of heterotrophic bacterial population along successional
sequence on spoil of brown coal colliery substrate, Folia Microbiol. 50 (2005)
427e435.
[65] C.J.F. ter Braak, P. 
Smilauer, CANOCO Reference Manual and CanoDraw for
Windows User's Guide, Software for Canonical Community Ordination,
Microcomputer Power, Ithaca, New York, 2002. Version 4.5.
[66] S. Scheu, Analysis of the microbial nutrient status in soil microcompartments:
earthworm faeces from a basalt-limestone gradient, Geoderma 56 (1993)
575e586.
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e5554

[67] B.A. Byzov, N.V. Khomyakov, S.A. Karin, A.V. Kurakov, Fate of soil bacteria and
fungi in the gut of earthworms, Eur. J. Soil Biol. 43 (2007) 149e156.
[68] K. Ravasz, L. T
oth, Studies on the gut actinomycete population of Eisenia fetida
(Savigny) (Oligochaeta: Lumbricidae), in: R. L
esel (Ed.), Microbiology in Poe-
cilotherms, Elsevier Science Publishers, 1990, pp. 141e144. B. V. (Biomedical
Division).
[69] C.A. Edwards, K.E. Fletcher, Interactions between earthworms and microor-
ganisms in organic-matter breakdown, Agr. Ecosyst. Environ. 24 (1988)
235e247.
[70] M.A. Horn, A. Schramm, H.L. Drake, The earthworm gut: an ideal habitat for
ingested N
2
O-producing microorganisms,, Appl. Environ. Microbiol. 69 (2003)
1662e1669.
[71] J. Ihssen, M.A. Horn, C. Matthies, A. Gr€
oßner, A. Schramm, H.L. Drake, N
2
O-
producing microorganisms in the gut of the earthworm Aporrectodea cal-
iginosa are indicative of ingested soil bacteria, Appl. Environ. Microbiol. 69
(2003) 1655e1661.
[72] B.A. Knapp, J. Seeber, S.M. Podmirseg, E. Meyer, H. Insam, Application of
denaturing gradient gel electrophoresisfor analysing the gut microflora of
Lumbricus rubellus Hoffmeister under different feeding conditions, B. Entomol.
Res. 98 (2008) 271e279.
[73] K.-H. Shin, J. Chun, Ch.-J. Cha, I.S. Kim, H.G. Hur, Analysis of anaerobic bacterial
community in the earthworm (Eisenia fetida) intestine, Agric. Chem.
Biotechnol. 47 (2004) 147e152.
[74] K. Fischer, D. Hahn, W. Hoenerlage, J. Zeyer, Effect of passage through the gut
of the earthworm Lumbricus terrestris L. on Bacillus megaterium studied by
whole cell hybridization, Soil Biol. Biochem. 29 (1997) 1149e1152.
[75] P. Lavelle, Faunal activities and soil processes: adaptive strategies that
determine ecosystem function, Adv. Ecol. Res. 27 (1997) 93e132.
[76] K. Thummes, J. Sch€
afer, P. K€
ampfer, U. J€
ackel, Thermophilic methanogenic
Archaea in compost material: occurrence, persistence and possible mecha-
nisms for their distribution to other environments, Syst. Appl. Microbiol. 30
(2007) 634e643.
[77] N. Yamamoto, R. Asano, H. Yoshii, K. Otawa, Y. Nakai, Archaeal community
dynamics and detection of ammonia-oxidizing archaea during composting of
cattle manure using culture-independent DNA analysis, Appl. Microbiol. Biot.
90 (2011) 1501e1510.
[78] Å. Frostegård, E. Bååth, The use of phospholipid fatty acid analysis to estimate
bacterial and fungal biomass in soil, Biol. Fertil. Soils 22 (1996) 59e65.
[79] J.A. Erwin, Fatty acids in eukaryotic microorganisms, in: J.A. Erwin (Ed.), Lipids
and biomembranes of eukaryotic microorganisms, Academic Press, New York,
1973, pp. 41e143.
[80] D.A. Bossio, K.M. Scow, Impact of carbon and flooding on soil microbial
communities: phospholipid fatty acid profiles and substrate utilization pat-
terns, Microb. Ecol. 35 (1998) 265e278.
A. Koubov
a et al. / European Journal of Soil Biology 68 (2015) 42e55 55